Methods

Dissections

GENERAL DISSECTION

Eye

To anesthetize crustaceans for dissection, place them on ice for 25-45 minutes. To begin dissection, first remove all limbs and mouth parts from the animal. Next, use ‘crushers’ to break the shell around the outer edge, being careful not to get too close to the eyestalks. Use the scraper to gently separate the muscle from the inside of the shell. Break the shell away from the body in order to free the face of the crustacean. Use the scraper again to separate the muscle from the bottom shell, focusing on the area near the eyes and exposing the two thin gastric muscles just below the eyes. Then use large scissors to separate the eyestalks and surrounding shell from the rest of the crab (making sure to cut laterally). Place the eyestalks into crab saline and put them in a refrigerator until you are ready to dissect them.

Stomach

Using the forceps, gently hold the start of the esophagus, which can be found right under the eyestalks that were just removed. As you hold the esophagus in one hand, in the other use a small pair of scissors to separate the stomach from the rest of the crab, ending the dissection at the end of the pylorus. Once the stomach is removed, gently rinse it with crab saline and then use the scissors to cut through the stomach following the esophagus to the pylorus. On the end opposite of the esophagus, make two butterfly cuts diagonally from main cut in order to allow the stomach to lie flat. Using the small scissors, remove the three teeth from the stomach, and then pin it in a dissecting dish with the side that once had the teeth face down. Gently pour enough crab saline to cover the stomach and then place the dish in the fridge until dissection.

Pericardial Organ (P.O.)

After removing the stomach, use the scissors and the scraper to separate the tissue from the remaining shell, and then remove the shell altogether. With a large pair of scissors, cut off the rounded sides of the crab (both left and right) and remove the gills. Using a scraper, gently remove the rest of the crab parts left in the body cavity. Next, using a large pair of scissors cut down the center of the crab, separating the left from the right. Place both parts in saline and put them in the fridge until you are ready to dissect them.

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Crab dissection

Stomach fully isolated from crab

MICRODISSECTION

Eye

After removing the eyestalks from the fridge, use the scissors to cut down middle between the two eyes, separating them. Gently remove the excess shell from around both eyes, until they are free. Holding the eye in one hand, use a small pair of scissors to make an incision from the base of the eye stalk up through the retina. Make an identical incision on the opposite side of the eyestalk, and repeat for the other eye (note: each eyestalk should still be in one piece when you are finished). Place each eyestalk in a dish with crab saline for dissection and use a dissecting microscope for the remainder of the dissection. Under the microscope use a pair of dissecting forceps and scissors to remove the eye tissue from the outer shell. The sinus gland can be identified by its brighter white or sometimes opal color. Remove the majority of the unwanted tissue, making sure to keep some tissue surrounding the sinus gland and parts of the retina and optic nerve for orientation. It is important to change the saline every 15-20 minutes in order to keep the sample cool and remove excess tissue in the solution. When the dissection is complete, fill a new dish with crab saline. Using forceps, gently place the dissected eyestalk into the new dish and pin it down. Cover the dish, and using tape, label the dish with the date, genus, species and sex of the organism, and the type of tissue.

Stomach

Remove the stomach from the fridge and continue the dissection under the microscope. On the esophagus end, locate the brain of the crustacean. Gently follow the nervous tissue eradiating from the brain and using a pair of dissecting scissors separate the brain from the rest of the tissue making sure not to remove too much nervous tissue. Place the brain in a clean dish with crab saline, pin it down, cover it, and label it according to the previous system. Back at the stomach, near where the brain was removed, there should be more nervous tissue. Remove the commissural ganglion (COG), the oesophageal ganglion (OG), the stomatogastric ganglion (STG), and surrounding nervous tissue (see picture more detail). Place the stomatogastric nervous system (STNS) in a clean dish with saline, carefully pin it, then cover and label it.

P.O.

Remove one pericardial organ from the fridge, place it in a deep dissecting dish with enough crab saline to cover the entire thing and put it under the microscope. The P.O. is typically located in a small hole on the part of the tissue that would have faced the inside of the crab. With the part of the tissue where the limbs once were face down in the dish, the aforementioned hole should be in the lower center part of the tissue. Remove the pericardial organ (with as little tissue on it as possible) using dissecting forceps and scissors, making sure to include both nerve bars.

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Close-up view of crab eye

P.O., ready for microdissection

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A carefully pinned STNS

Fixing

After each dissection was completed the tissue was carefully pined as flat as possible into the fix dishes, covered with saline, and labeled in both in the master book and on the dish. All of the tissue samples collected during that day would then be fixed overnight. The fix solution consisted of 4% Paraformaldehyde, The entirety of the Paraformaldehyde ampoules’ (10ml, pre-diluted) was mixed with 30ml of PBST to total 40 ml of liquid. Using gloves, the saline was poured out of each of the fixing dishes and was replaced with the 4%PARA solution. The tissues would sit in the fix solution overnight. After the fixation process each tissue was rinsed 5 times every half hour using PBST. One half hour after completing the rinses, each tissue was assigned one of three test antibodies: a-CLDH (calcitonin like diuretic hormone), a-melatonin receptor, a-melatonin. For each of these antibodies a solution was made and placed into the Eppendorf tubes.

Antibodies

The primary antibodies were comprised of 10% Normal Donkey Serum (NDS), phosphate buffer solution with tricane-x (PBST), and antibody. The melatonin antibody was made with a ratio of 1:500, the melatonin receptor antibody was made with a ratio of 1:300, and the calcitonin-like anti-diuretic hormone (CLDH) was made at a ratio of 1:1000. These ratios represent the amount of given antibody to the total volume.

The solution for each antibody was then placed carefully into Eppendorf tubes, using extreme care to not cross contaminate. For eyestalks the tubes were filled halfway, while the other tissues (P.O and S.T.N.S) were filled all the way. Each tissue was next carefully removed from the fix dishes using microscopes to properly remove the pins and placed into the properly labeled Eppendorf tubes. The tissue then sat in the primary antibody for 3 days. Next a second rinse was conduced rinsing the tissue 5 times in PBST every half hour. One half hour after the last rinse the tissues were moved into the same secondary antibody a- donkey anti rabbit solution:

The secondary antibody was made in a similar fashion to the primary antibodies. We used a 1:300 ratio of antibody to total volume, again with 10% comprised of NDS, antibody, and the remaining volume of PBST. As mentioned, regardless of the primary antibody used, all tissues received the same secondary antibody.

The secondary antibody was extremely sensitive to light, and both the Eppendorf tubes and the solutions need to be covered with tinfoil at all times possible. The tissues would sit in the secondary antibody overnight. In the morning a final rinse was conducted, 5 rinses, every 30 minutes in PBS and one hour after the final rinse the tissue was removed from the Eppendorf tubes and mounted.

Mounting

Slides were prepared using 2-3 drops of Vectashield with or without DAPI. Tissue samples were added and covered with a clear glass cover slip sealed with clear nail polish. They were labeled according to their tissue type, species, and primary and secondary antibodies.

Microscopy

Fluorescence and brightfield microscopy After mounting, we waited an appropriate amount of time for the slides to dry. All slides were initially looked at under a Zeiss Axiovert 200 fluorescence microscope. We viewed a total of 59 stained slides under the fluorescent microscope and chose selected slides to be imaged in more detail using an Olympus Fluoview 1000 confocal microscope.